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Biochimie 110 (2015) 52e61

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Biochimie

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Research paper

Activation of p53 mediated glycolytic inhibition-oxidative stress-apoptosis pathway in Dalton's lymphoma by a ruthenium (II)-complexcontaining 4-carboxy N-ethylbenzamide

Raj Kumar Koiri a, Surendra Kumar Trigun b, *, Lallan Mishra c

a Department of Zoology, Dr. Harisingh Gour Central University, Sagar, Madhya Pradesh 470003, Indiab Biochemistry Section, Department of Zoology, Banaras Hindu University, Varanasi, Uttar Pradesh 221005, Indiac Department of Chemistry, Banaras Hindu University, Varanasi, Uttar Padesh 221005, India

a r t i c l e i n f o

Article history:Received 25 September 2014Accepted 30 December 2014Available online 8 January 2015

Keywords:Rutheniump53PFKFB3LDHApoptotic factors

Abbreviations: Bcl-2, B-cell leukemia/lymphoma-2fructose-2,6-bisphosphate; GPx, glutathione peroxidaiPFK2/PFKFB3, inducible isoform of 6-phosphofructo-drogenase; OXPHOS, oxidative phosphorylation; RORu(II)-CNEB, [Ru(CNEB-H)2(bpy)2] 2PF6$0.5 NH4PF6 (taining 4-carboxy N-ethylbenzamide as ligand); SOD,* Corresponding author. Tel.: þ91 9415811962.

E-mail address: sktrigun@gmail.com (S.K. Trigun).

http://dx.doi.org/10.1016/j.biochi.2014.12.0210300-9084/© 2015 Elsevier B.V. and Soci�et�e française

a b s t r a c t

There is a general agreement that most of the cancer cells switch over to aerobic glycolysis (Warburgeffect) and upregulate antioxidant enzymes to prevent oxidative stress induced apoptosis. Thus, there isan evolving view to target these metabolic alterations by novel anticancer agents to restrict tumorprogression in vivo. Previously we have reported that when a non toxic dose (10 mg/kg bw i.p.) of a novelanticancer ruthenium(II)-complex containing 4-carboxy N-ethylbenzamide; Ru(II)-CNEB, was adminis-tered to the Dalton's lymphoma (DL) bearing mice, it regressed DL growth by inducing apoptosis in theDL cells. It also inactivated M4-LDH (M4-lactate dehydrogenase), an enzyme that drives anaerobicglycolysis in the tumor cells. In the present study we have investigated whether this compound is able tomodulate regulation of glycolytic inhibition-apoptosis pathway in the DL cells in vivo. We observed thatRu(II)-CNEB could decline expression of the inducible form of 6-phosphofructo-2-kinase (iPFK2:PFKFB3), the master regulator of glycolysis in the DL cells. The complex also activated superoxide dis-mutase (the H2O2 producing enzyme) but declined the levels of catalase and glutathione peroxidase (thetwo H2O2 degrading enzymes) to impose oxidative stress in the DL cells. This was consistent with theenhanced p53 level, decline in Bcl2/Bax ratio and activation of caspase 9 in those DL cells. The findingssuggest that Ru(II)-CNEB is able to activate oxidative stress-apoptosis pathway via p53 (a tumorsupressor protein) mediated repression of iPFK2, a key glycolytic regulator, in the DL cells in vivo.

© 2015 Elsevier B.V. and Soci�et�e française de biochimie et biologie Mol�eculaire (SFBBM). All rightsreserved.

1. Introduction

Identification of cellular/molecular targets is of prime concernfor formulating novel anticancer agents [1]. Due to effective bio-distribution and multimodal cellular actions, during recent past,ruthenium complexes have drawn much attention as next gener-ation anticancer agents [2]. So far mechanistic aspects of

; DL, dalton's lymphoma; FBP,se; GSH, glutathione reduced;2-kinase; LDH, lactate dehy-S, reactive oxygen species;Ruthenium(II)-complex con-superoxide dismutase.

de biochimie et biologie Mol�ecula

anticancer metal complexes are concerned, DNA, once consideredas their main target [3,4], is now evident to be highly unselective[5] and therefore, there is an evolving concept to evaluate whethermetal complexes could be able to attenuate certain tumor growthassociated biochemical events at cellular level [5e7]. In thisrespect, Ru-complexes get an upper hand, as metal center ofruthenium has been shown to interact with a variety of ligands [8]and thereby enabling these complexes to affect various cellularactivities [9].

We have synthesized and characterized a ruthenium complex;Ru(II)-CNEB, which was found to be highly biocompatible to micewhen administered in vivo. Additionally, it could interact with andinhibit M4-LDH non-competitively both in vitro and at tissue level[10]. During pilot experiments, though this compound did not showany DNA cleavage activity in vitro (Supplementary data with thisarticle), when administered to the DL bearing mice, it produced

ire (SFBBM). All rights reserved.

R.K. Koiri et al. / Biochimie 110 (2015) 52e61 53

apoptotic pattern of DNA cleavage which was consistent with thedecreased DL cell viability and increased life span of the tumorbearing mice [11]. Importantly, since, this complex increasedcertain apoptotic markers and also inhibited M4-LDH (LDH-5) inthose DL cells, it was argued that this complex, instead of targetingDNA directly, is able to activate glycolytic inhibition-apoptosispathway in a tumor cell in vivo [11]. This necessitated furtherstudies on characterizing regulatory factors of this pathway astarget of Ru (II)-CNEB in the DL cells in vivo.

Switching over to aerobic glycolysis, known as ‘Warburg effect’,is considered to be a common trait of most of the growing tumors[12] and therefore, inhibition of enhanced tumor glycolysis is nowadvocated as one of the therapeutic strategies in cancer therapy[13,14]. In this regard, targeting regulatory enzymes of glycolyticpathway assumes special importance for the novel anticanceragents. Phosphofructokinase1 (PFK1) catalyzes committed step ofglycolysis. PFK2 domain of D-fructose-6-phosphate-2-kinase/fruc-tose-2,6-bisphosphatase (PFK2/FBPase2) synthesizes fructose-2,6-bisphosphate (FBP) to activate PFK1 and thereby, it acts as mainglycolytic regulator under a variety of pathological challengesincluding tumor progression [15]. Cancer cells express C type PFK1which is more sensitive to FBP [16] and also over express a cata-lyticallymore efficient inducible form of PFK2 (iPFK2: PFKFB3 gene)[17]. Since, iPFK2 repression has been reported to inhibit tumor cellgrowth in vitro [13], this glycolytic activator could be considered asrelevant therpautic target for the novel anticancer agents. Similarly,to sustain enhanced glycolysis, tumor cells adapt to produce lactatefrom pyruvate by activating LDH-5. Reports also suggest that overexpression of LDH-5 gene (LDH-A) is associated with tumor growth[12,18] and thus, advocating repression of LDH-5 gene as anothermechanism to define therapeutic target for a novel anticanceragent.

So far up stream regulation of the cell bioenergetics is con-cerned, p53, a tumor suppressor protein, has been found tomodulate overall balance between glycolysis and mito-OXPHOS[19]. The loss of normal p53 has been reported to be criticallyassociated with tumor progression, particularly in leukemia andlymphoma [20]. Similarly, p53 over expression has been demon-strated to have prognostic significance in lymphoma [21]. Recently,p53 has been reported to regulate tumor cell energy metabolism[22] via down regulating expression of glucose transporters GLUT1and GLUT4 [23] and also by activating the TIGAR gene which re-presses iPFK2/PFKFB3 [24]. Thus, when tumor cells switch over toglycolytic phenotype, it is accompanied with declined p53expression [25]. Similarly, inhibition of glycolysis has been shownto activate p53 [26].

It has been suggested that ROS are down stream mediators ofp53 dependent apoptosis [27], wherein, Bcl-2/Bax ratio acts asmain determinant of this pathway [28]. Low level of p53 inducesexpression of antioxidant enzymes responsible to maintain ROSlevel within a permissive range which otherwise can cause DNAdamage and genomic instability in the cells [29]. Higher level of p53protein, on the other hand, is known to enhance the expression ofpro-oxidant and proapoptotic factors [30].

Tumor cells, in addition to acquiring glycolytic phenotype, alsomodulate their oxygen metabolism in many ways [12,31]. Super-oxide dismutase (SOD), catalase, glutathione peroxidase (GPx) andglutathione reductase (GR) constitute main antioxidant defensemechanism in most of the cells. The oxygen free redicals (O2

.-),produced during mitochondrial oxidative phosphorylation, aredismutated to hydrogen peroxide (H2O2) by SOD followed by con-version of H2O2 into water by catalase and GPx [32]. As SOD is thecommitted enzyme of this pathway responsible to produce H2O2, itappears to be the most relevant target for therapeutic interventionin the tumor cells. In the recent past, both SOD isoforms (CueZn-

SOD: cytosolic; SOD1 and Mn-SOD: mitochondrial; SOD2) havebeen found to be implicated in tumerogenesis [33].

Some earlier reports have described that reduced levels of SOD1& SOD2 facilitate tumerogenesis [34,35] via maintaining low levelof H2O2 in the cancerous cells and accordingly, increased SOD ac-tivity and in turn, higher level of H2O2 may contribute for tumorsuppression [36,37]. However, effectiveness of this SOD mediatedmechanism depends on the status of the two down stream en-zymes; catalase and GPx in the cancerous cells. Indeed, SOD andcatalase double transfectant (SOCAT3) cells and cells with overactivated GPx1 have shown protection from oxidative cell damage[38,39]. Thus, to characterize oxidative mechanism based anti-cancer potential of a novel anticancer agent, it is important to studycomparative profile of the three antioxidant enzymes involved inH2O2 metabolism in the cancer cells.

In the present article, we have investigatedwhether Ru(II)-CNEBis able to activate p53 mediated glycolytic inhibition-oxidativestress-apoptosis pathway in the DL cells in vivo.

2. Materials and methods

2.1. Chemicals

Ruthenium(II)-complex containing 4-carboxy N-ethyl-benzamide as ligand, Ru(II)-CNEB; [Ru(CNEB-H)2(bpy)2] 2PF6$0.5NH4PF6, whose structural details have already been describedpreviously [10,11], was used in the present study. Antibodiesagainst p53, caspase 9, Bcl-2, Bax, SOD1, SOD2 & PFK 2 werepurchased from Santa Cruz and b-actin was purchased fromSigmaeAldrich Co., USA. HRP- conjugated anti rabbit/goat/mouseIgG were obtained from Genei. 20,70-dichlorofluorescin diacetateand ECL super signal western pico kit were purchased from Flukaand Pierce respectively. Lactate estimation kit was purchasedfrom Biorex Diagnostics, Ltd, UK. Other general chemicals andreagents were obtained from Merck or SRL unless otherwisespecified.

2.2. Induction of Dalton's lymphoma (DL) in mice

Inbred AKR strain mice of 16e18 weeks age weighing 24e26 gwere used for the experiments. Mice were maintained at standardlaboratory conditions with the supply of food and water ad libitum.This work was approved by the institutional animal ethical com-mittee (Dan/2006-07/962). Dalton's lymphoma was induced bytransplantation of 1 � 107 viable tumor cells (assayed by trypanblue method) i.p. per mice. Development of DL was confirmed byabnormal belly swelling and increased body weight which becamevisible on 10e12th post transplantation day. The DL bearing micesurvived up to 18 ± 2 days.

2.3. Treatment schedule

The DL mice were randomly divided into 2 groups with 10 micein each. The first group DL mice were treated with Ru(II)-CNEBcomplex (10 mg/kg bw/day, ip), and the second group, designatedas DL control, were similarly injected with equal volume of KrebsRinger Buffer (KRB). As DL becomes visible on day 10e11 and DLbearing mice survived up to 18e20 days post DL transplantation,the treatments with the compounds were started from day 11 oftumor transplantation and continued up to day 17th. The normalcontrol group mice were also treated simultaneously with KRB. Tostudy biochemical/molecular parameters, 3-4 mice from eachgroup were sacrificed on day 18th and tumor ascites pooled from 3to 4 DL mice from each group were centrifuged at 2000 � g at 4 �Cto collect DL cells. The DL cell extract was prepared using lysis

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buffer (20 mM Tris-Cl, pH 7.4, 0.15 M NaCl, 1 mM EDTA,1 mM EGTA,1% Triton X-100, 25 mM Na2 pyrophosphate and 1 mM PMSF). Thecell lysates were centrifuged at 20,000 � g for 30 min and super-natant obtained were used for biochemical and molecular studies.Protein concentrations in the extracts weremeasured following themethod of Lowry et al. [40].

2.4. Western blotting

For western blot analysis of different proteins, DL cell extractscontaining 60 mg proteins, were subjected to 10% SDS-PAGE. Asdescribed previously [11], proteins were transferred to nitrocellu-lose membrane followed by detection of p53, caspase 9, Bcl-2, Bax,iPFK2, SOD1 and SOD2 against specific polyclonal antibodies(1:1000). Proteins on membrane were detected by ECL west picokit. As loading control, b-actin was probed similarly using mono-clonal anti-b-actin-peroxidase antibody (1:10,000). Protein bandswere quantified using gel densitometry software AlphaImager2200.

2.5. Assay of the antioxidant enzymes

Catalase activity was measured following an earlier method [41]with somemodifications. Briefly, 1 ml reactionmixture consisted of0.05 M phosphate buffer (pH 7.0) and 0.003% H2O2. The reactionwas started by the addition of suitably diluted cell extract anddecrease in absorbance at 240 nmwas recorded for 10 min. Unit ofthe enzymewas defined as mmol of H2O2 depleted/min. The activitywas expressed as units/mg protein.

Glutathione peroxidase (GPx) activity was determined asdescribed earlier [42] with slight modifications. Briefly, cell ex-tracts were added to the assay buffer consisting of 100 mM Tris-Cl (pH 7.2), 3 mM EDTA, 1 mM sodium azide, 0.25 mM H2O2,0.5 mM NADPH, 0.17 mM GSH, and 1 unit GR. The change inabsorbance per minute at 340 nm was recorded for 10 min andenzyme activity was expressed as mmole NADPH oxidized/min/mg protein.

2.6. Analysis of SOD and Gpx by non-denaturing PAGE

The active levels of superoxide dismutase (SOD) and glutathioneperoxidase (GPx) was determined using non-denaturing PAGE ofthe cell extracts following the method recently reported from ourlab [42,43]. The cell extract containing 60 mg proteinwere loaded ineach lane of 10% non-denaturing PAGE. After electrophoresis at4 ± 2 �C, the gels were subjected to substrate specific staining ofdifferent antioxidant enzymes.

The staining mixture for SOD consisted of 2.5 mM NBT, 28 mMriboflavin, and 28 mM TEMED. Gels were incubated for 20 min inthe dark, followed by exposing them to a fluorescent light tillachromatic SOD bands developed against dark blue background.

GPx specific staining mixture was composed of 50 mM TriseClbuffer (pH 7.9), 3 mM GSH, 0.004% H2O2, 1.2 mM NBT and 1.6 mMPMS. Achromatic bands corresponding to GPx activity appearedagainst a violeteblue background.

The intensities of all the bands were quantified by gel densi-tometry using AlphaImager 2200 gel documentation software.Specificity of the PAGE bands of different enzymes were confirmedby obtaining clear negative results when similarly run gels weretreated in the absence of the enzyme specific substrates. In eachcase, PAGE was performed at least 3 times and mean ± SD ofdensitometry values of the bands, as percentage of the control lane,have been presented in the result with one representative gelphotograph.

2.7. Biochemical estimations

2.7.1. LactateThe concentration of lactate was determined by measuring

lactate oxidation by lactate oxidase as per the manufacturer's in-structions given in the lactate assay kit obtained from Biorex Di-agnostics Ltd, UK.

2.7.2. H2O2

Following the method described earlier [42]; intracellular H2O2was determined by measuring H2O2 dependent oxidation of DCFH-DA into DCF. Briefly, 100 ml of cell extracts were incubated with10 mM DCFH-DA at 37 �C for 30 min in dark. Using excitation at504 nm and emission at 529 nm, intensity of DCF fluorescence wasmeasured. H2O2 concentration in the extract was determined byusing a calibration plot made against different H2O2 concentrations(5e100 mM).

2.7.3. Total glutathione (GSH þ GSSG)Following the method described earlier [42], DL cell extracts

were precipitated with 5% sulfosalicylic acid in the ratio of 1:2 andcentrifuged. The supernatant collected was neutralized. In a 96 wellmicro plate, 50 ml neutralized supernatant was incubated with100 ml of the reagent containing 0.30 mM NADPH, 0.22 mM DTNBand 1.6 units/ml GR prepared in 100 mM phosphate buffer (pH 7.4)containing 1mM EDTA. The absorbance was recorded at 412 nm for10 min using Micro Scan MS5608A (ECIL) micro plate reader. Totalglutathione content was expressed in terms of nmol/mg protein.

2.8. Semi-quantitative RT-PCR

Total RNA was isolated from DL cells using TRI reagentfollowing the manufacturer's protocol. DNA-free™ (Ambion) wasused to remove any contaminating DNA from the RNA preparationfollowing the manufacturer's protocol. Briefly, reaction mixtureconsisting of DNase I, 10� DNase I buffer and RNA sample wasincubated at 37 �C for 20 min. Thereafter, DNase I inactivationreagent slurry was added and incubated for 2 more min at roomtemperature and centrifuged. The upper phase was collected asDNA free RNA solution. From this RNA isolate, 2 mg RNA wassubjected for reverse transcription using 200 U of reverse tran-scriptase and 200 ng random hexamer to make ss-cDNA (RevertAid First strand cDNA synthesis kit, MBI fermentas). The PCR re-action mixture contained 1� Taq polymerase buffer, 0.2 mM eachof the four dNTPs, 1.0 U of Taq polymerase, and 10 pmol of thespecific primer. The mouse gene-specific primers used were: Bcl-2(forward 50-TAC CGT CGT GAC TTC GCA GAG-30; reverse 50-GGCAGG CTG AGC AGG GTC TT-30); Bax (forward 50-CGG CGA ATT GGAGAT GAA CTG-30; reverse 50-GCA AAG TAG AAG AGG GCA ACC-30);PFKFB3 (forward 50-GGC AAG ATT GGG GGC GAC TC-30; reverse 50-GGC TCC AGG CGT TGG ACA AG-30); LDH A (forward 50-ATG CACCCG CCT AAG GTT CTT-30; reverse 50-TGC CTA CGA GGT GAT CAAGCT-30); SOD2 (forward 50-GCA CAT TAA CGC GCA GAT CA-30;reverse 50-AGC CTC CAG CAA CTC TCC TT-30) and b actin (forward50-ATC GTG GGC CGC TCT AGG CAC C-30; reverse 50-CTC TTT GATGTC ACG ATT TC-30). Linearity of PCR amplifications was checkedfor each gene using various cycles (20, 24, 28, 32, 36 and 40 cycles)vs densitometric value of the PCR product of the correspondinggene. Accordingly, the optimal number of cycles from the linearphase and other conditions were chosen for amplification for eachgene as; b actin, 30 cycles of 30 s at 94 �C, 30 s at 52 �C, and 30 s at72 �C; Bcl-2 & LDH A, 34 cycles of 45 s at 94 �C, 45 s at 55 �C, and1 min at 72 �C; Bax, 30 cycles of 30 s at 94 �C; 60 s at 56 �C, and1 min at 72 �C; PFKFB3, 31 cycles of 60 s at 95 �C, 60 s at 50 �C, and1 min at 72 �C; SOD2, 27 cycles. The amplification products,

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analyzed by 1e2% agarose gel electrophoresis, were visualized byethidium bromide staining and were found to be; Bcl 2 (350 bp),Bax (160 bp), PFKFB3 (329 bp), LDH A (103 bp), SOD2 (241 bp) andb actin (543 bp). Amplification of b actin served as a control. ThreeRT-PCR repeats from 3 RNA isolates for each genewere performed.The expression levels were measured by densitometry usingAlphaImager 2200.

2.9. Statistical analysis

Experimental data were expressed as mean ± SD and Student's ttest was applied for determining the level of significance betweencontrol and experimental groups and value of p < 0.05 wasconsidered significant.

Fig. 1. Effect of Ru(II)-CNEB (Rc) on the expression of LDH-A and iPFK2 & lactate productionratio of LDH-A/b actin mRNA presented as mean ± SD from 3 RT-PCR repeats. (b) showsphotograph for PFKFB3 with the ratio of PFKFB3/b actin mRNA presented as mean ± SDprotein in each lane, presented with the relative densitometric values of iPFK2/b actin as meDL groups).

3. Results

3.1. Ru(II)-CNEB mediated decline of lactate level in the DL cellsin vivo

Lactate serves as an alternate metabolic fuel for the high energydemanding tumor cells. Therefore, overproduction of lactate isconsidered one of the hallmarks of the tumor growth. According toFig. 1b, as compared to the DL cells from untreated group, there is asignificant decline (p < 0.001) in lactate level in case of the DL cellsfrom Ru(II)-CNEB treated DL mice. To ascertain, whether decline inlactate level is due to decline in the expression of M4-LDH, the levelof LDH-A mRNAwas measured in the DL cells from the untreated &the treated group DL mice. According to Fig. 1a, the level of LDH-A

in the DL cells in vivo. (a) shows representative RT-PCR photograph for LDH-A with thelactate level represent as mean ± SD where n ¼ 4. (c) shows representative RT-PCRfrom 3 RT-PCR repeats. In (d), a representative western blot photograph, with 60 mgan ± SD from three western blot repeats. *p < 0.05; ***p < 0.001 (untreated vs. treated

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mRNA in the DL cells from Ru(II)-CNEB treated DL mice is found tobe unchanged as compared to the untreated counterpart.

3.2. Downregulation of iPFK2/PFKFB3 in the DL cells due to thetreatment with Ru(II)-CNEB

Over expression of iPFK2/PFKFB3 is associated with tumorgrowth. Recently, p53 has been reported to regulate tumor cellenergy metabolism indirectly by activation of the TIGAR genewhich declines iPFK2/PFKFB3 induced fructose-2,6-bisphosphateproduction in the tumor cells. As illustrated in Fig. 1c and d, thelevels of PFKFB3 mRNA and its protein product declined signifi-cantly (p < 0.05e0.001) in the DL cells from Ru(II)-CNEB treated DLmice than that from the untreated counterpart.

3.3. Ru(II)-CNEB increases p53 level in the DL cells in vivo

Restoration of p53 level is an important strategy of anticancerchemotherapy. Based on immunoblot analysis, level of p53 wasobserved to be ~5 times higher in the DL cells (p < 0.001) from theRu(II)-CNEB treated DL mice than that from the control group mice(Fig. 2).

3.4. Enhanced expressions of SOD1 & SOD2 in the DL cells fromRu(II)-CNEB treated DL mice

Oxidative stress and mitochondrial dysfunction are known toinitiate final steps of apoptosis. Infact ROS has been suggested to actas a down streammediator of p53 dependent apoptosis. SOD is thefirst commited enzyme that neutralizes superoxide anion freeradical (O2

��) based oxidative stress in the cells. According to Fig. 3a,the level of active SOD2 & SOD1, both shows significant incrementsin the DL cells from Ru(II)-CNEB treated DL mice. To confirmwhether activity increment is contributed due to the similar in-crease in the expression of these antioxidant enzymes, RT-PCR andwestern blot analysis was performed. Fig. 3bed illustrate thatRu(II)-CNEB is able to increase the expression of both, SOD1 andSOD2 significantly (p < 0.05e0.001) in the DL cells in vivo.

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Fig. 2. Treatment with Ru(II)-CNEB (Rc) increased the level of p53 in the DL cellsin vivo. A representative western blot photograph is presented with the relativedensitometric values of p53/b actin as mean ± SD from three western blot repeats.***p < 0.001 (untreated vs. treated DL groups).

3.5. Ru(II)-CNEB mediated in vivo modulation of pro-oxiativefactors in the DL cells

The two antioxidant enzymes, catalase & GPx, down stream toSOD, show significant decline in their activity (p < 0.05) in the DLcells from Ru(II)-CNEB treated DL mice (Fig. 4aec). However,treatment with Ru(II)-CNEB caused a significant decrease in theactivity of GPx1, with a concomitant increase in the activity of GPx2(Fig. 4c). Such a reciprocal change between SODs and catalase &GPx is likely to allow accumulation of H2O2 at cellular level. Fig. 5ashows that indeed there is a marked increase (~6 times; p < 0.001)in H2O2 level in the DL cells from Ru(II)-CNEB treated DL mice thanthose from the untreated groups. Level of glutathione is anothermarker of oxidative stress in the cells. According to Fig. 5b, ascompared to the untreated DL group, the DL cells from Ru(II)-CNEBtreated DL mice show a significant decline (p < 0.05) in totalglutathione (GSH þ GSSG) level.

3.6. Ru(II)-CNEB mediated in vivo modulation of pro-apoptoticfactors in the DL cells

Bcl2/Bax ratio serves as one of the critical determinants ofapoptotic induction at cellular level. As compared to the untreatedDL mice, treatment with Ru(II)-CNEB caused a significant decline(p < 0.001) in the level of Bcl2 protein in the DL cells with aconcomitant increase in the Bax protein (Fig 6b). Such reciprocalchanges in Bcl2 vs Bax resulted into a significantly declined Bcl2/Bax ratio (~3.5 times; p < 0.01) in the DL cells from Ru(II)-CNEBtreated DL mice (Fig. 6b). The RT-PCR results (Fig 6a) also demon-strate a significant decline in Bcl2/BaxmRNA level, however, mainlydue to a significant decrease in Bcl2 mRNA level in the DL cells fromthe treated rats.

Caspase 9 cleavage is an important event of the mitochondrialpathway of apoptosis and that of PARP-1 cleavage associates withDNA fragmentation in the apoptotic cells. As shown in Fig. 6c,treatment with Ru(II)-CNEB is able to produce cleaved products ofcaspase 9. Full length PARP-1 is a 116 kDa protein, which, uponactivation, is cleaved into a 89 kDa C-terminal & a 24 kDa N-ter-minal protein fragments. Fig 6d illustrates ~1.5 times increase in thelevels of the cleaved PARP-1 fragments with a proportionatedecline in its full length PARP-1. This was consistent with the DNAfragmentation pattern of the DL cells reported previously [11].

4. Discussion

The two main findings reported in our previous paper [11]; one,Ru(II)-CNEB could inhibit M4-LDH and second, it induced release ofmitochondrial cytochrome c in the DL cells in vivo, hinted towardsinduction of metabolic derangement led apoptosis in the DL cellsdue to the treatment with this compound. This tempted us toexplore the regulatory aspects of such cellular effects produced byRu(II)-CNEB. Since, decline inM4-LDH activity in the DL cells, due tothe treatment with an anti-DL agent, could be attributed to thereduced expression of this enzyme [28], first we checked the mRNAlevel of LDH-A (M4-LDH) in the DL cells from Ru(II)-CNEB treatedDL mice. The RT-PCR result (Fig. 1a) shows no change in the level ofLDH-AmRNA in the DL cells due to the treatment with Ru(II)-CNEB.Thus, suggesting that Ru(II)-CNEB does not affect expression of M4-LDH. This implies that decline in M4-LDH activity, observed pre-viously due to the treatment with Ru(II)-CNEB [11], is a conse-quence of inhibiting catalytic efficiency of this LDH isozyme by thecompound at protein level. Indeed, this complex has already beendemonstrated to interact with and inhibit M4-LDH non-competitively [10].

Fig. 3. Effect of Ru(II)-CNEB (Rc) on (a) active levels of SOD2 & SOD1, (b) on the expression of SOD1 and (c & d) on the expression of SOD 2 in the DL cells from untreated and treatedDL mice. (a) gel photograph is a representative of 10% native PAGE of 60 mg protein in each lane, presented with the relative densitometric values for the respective SOD bands from3 PAGE repeats. (c) shows a representative RT-PCR photograph with the densitometric values of SOD2/b actin mRNA presented as mean ± SD from 3 RT-PCR repeats. (b and d) showsrepresentative western blot photographs, with 60 mg protein in each lane, presented with the densitometric values of SOD1/b actin and SOD2/b actin respectively as mean ± SD from3 western blot repeats. *p < 0.05; **p < 0.01; ***p < 0.001 (untreated vs. treated DL groups).

R.K. Koiri et al. / Biochimie 110 (2015) 52e61 57

Overproduction of lactate is considered as one of the hallmarksof the tumor growth [44]. It has been reported that tumor cellssurvival is greatly supported by excess of lactate produced by thetumor cells [45]. Also, blockage of tumor M4-LDH, responsible tosynthesize lactate, has been found to suppress this additional routeof metabolic supplementation and thereby renders tumor cellssusceptible to death [45]. Therefore, keeping aside the mechanismby which Ru(II)-CNEB decreases M4-LDH activity, the resultantdecline in lactate production could be of high therapeutic relevancefor this compound. Indeed, Ru(II)-CNEB, earlier found to inhibitM4-LDH [11], is also able to significantly decline lactate level in the

DL cells (Fig. 1b), and thus, advocating M4-LDH as a therapeutictarget for this compound at least in case of the DL cells in vivo. Sucha mechanism gets support from inhibition of tumor cell prolifera-tion in vitro due to the treatment with certain N-hydroxyindole-based inhibitors of M4-LDH [46].

Moreover, M4-LDH catalyzed rapid production of lactate ismainly dependent on the adequate supply of pyruvate as a conse-quence of enhanced glycolysis in the tumor cells [18,47]. Thecommitted step of glycolysis is catalyzed by PFK1 which is evidentto be the target of multimodal regulation under a variety of path-ophysiological conditions including tumor development [16]. PFK2,

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Fig. 4. Effect of Ru(II)-CNEB (Rc) on the activity of catalase (a) and GPx (b and c) in the DL cells in vivo. The values in (a and b) are mean ± SD, where n ¼ 4. (c) represents active levelof GPx in the DL cells from untreated and complex treated DL mice. The gel photograph is a representative of 10% native PAGE of 60 mg protein in each lane, presented with therelative densitometric values for the respective GPx bands from 3 PAGE repeats. *p < 0.05; **p < 0.01 (untreated vs. treated DL groups).

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the kinase domain of a bi-functional enzyme (PFK2/FBPase2),synthesizes FBP, the most effective allosteric activator of PFK1, andthereby, considered as a master regulator of the glycolytic pathway.In general, tumor cells are evident to express a C-type PFK1 thatshows greater sensitivity for FBP activation [16] and also express acatalytically more efficient isoform of PFK2 (iPFK2: PFKFB3) [13,17].Importantly, iPFK2 is found to be over expressed as a universal traitof most of the growing tumor cells including DL [28] and manytumors of human origin as well [17]. It was interesting to observethat this compound could significantly decline the expression ofiPFK2, both at mRNA and at protein levels (Fig. 1c and d). Thisclearly suggested that Ru(II)-CNEB is able to downregulate thesynthesis of master regulator of the glycolytic pathway in the DLcells in vivo. The argument is well supported by a report describingcorrelation between PFKFB3 gene silencing by siRNA and apoptosisin the Hela cells in culture [13]. Though information is scanty onmodulation of iPFK2 by synthetic compounds in tumor cells, a 3-(3-pyridinyl)-1-(4-pyridinyl)-2-propen-1-one has been found toinhibit iPFK2 resulting into decreased glucose uptake by the tumorcells leading into tumor growth suppression in vivo [48]. In thiscontext, the findings of Fig. 1c and d are first of its kind todemonstrate that Ru(II)-CNEB is able to repress iPFK2 in a tumorcell in vivo and thereby speculated to render less production ofadequate glycolytic intermediates to sustain high glycolytic effi-ciency of the DL cells in vivo.

So far tumor growth associated regulator of cell bioenergetics isconcerned, p53, a tumor suppressor protein, has been given muchemphasis as it has been reported to be involved in imposing War-burg effect during tumor development [19,22]. It is now evidentthat switching over to glycolytic phenotype by the tumor cells isaccompanied with the declined p53 level [25]. Similarly, theenhanced level of p53 has been found to repress glucose trans-porters GLUT1 and GLUT4 [23]. Also, increased p53 has beendemonstrated to decline the expression of iPFK2 via activating theTIGAR gene [24] and consequently, it inhibits glycolysis [26]. Wehave observed significant increase in p53 level (Fig. 2) vis a vis asignificantly declined iPFK2 expression (Fig.1c and d) in the DL cells

due to the treatment with Ru(II)-CNEB and thus suggesting an as-sociation between Ru(II)-CNEB mediated enhanced p53 level anddeclined activity of the committed step of glycolytic pathway in theDL cells in vivo.

It is known that activation of aerobic glycolysis by the tumor cellsis a metabolic strategy to prevent production of ROS, an inevitableoutcome of the oxidative energymetabolism, and thereby to protecttumor cells from ROS induced apoptosis [49]. In addition, tumorcells are known to modulate main antioxidant enzymes to preventoxidative stress [34,35]. Therefore, deranging such enzymaticmechanisms by a therapeutic agent is argued to be a relevant optionfor driving tumor cells to undergo apoptosis [28,43].

SOD is the first and commited enzyme of antioxidant pathwaythat neutralizes O2

�� into H2O2. It has been reported that reducedlevels of SOD1 and SOD2 maintain low level of H2O2 in thecancerous cells to facilitate tumerogenesis [34,35]. Similarly,increased SOD activity and in turn, higher level of H2O2 is specu-lated to inhibit tumor progression [36,37]. According to Fig. 3a,there is a significant increase in the activity of both the SOD iso-forms; SOD2 & SOD1, in the DL cells from Ru(II)-CNEB treated DLmice. Since, the pattern of active levels of both of them coincidedwith the similar increments in expression of both these enzymes(Fig. 3bed), it is evident that Ru(II)-CNEB is able to enhanceexpression of SOD2 and SOD1 to make overall increment in SODactivity in the DL cells in vivo. Since reports are limited on metalcomplex induced expression of antioxidant enzymes, it is a firstreport wherein, a Ru(II)-complex is demonstrated to enhance ac-tivity of SOD by overexpressing SOD2 and SOD1 proteins. Moreover,keeping aside these explanations, the enhanced level of SODs islikely to ultimately produce higher level of H2O2 in the DL cells dueto the treatment with Ru(II)-CNEB.

H2O2 produced by SODs is metabolized by GPx and Catalase andactivity of both these enzymes were observed to be declinedsignificantly in the DL cells from Ru(II)-CNEB treated DL mice(Fig. 4aec). Such a reciprocal pattern between SOD vs catalase &GPx has been reported accountable for unusually increased level ofH2O2 in the DL cells due to the treatment with emodin [50]. Indeed,

Fig. 5. Effect of Ru(II)-CNEB (Rc) on the level of H2O2 (a) and total glutathione (b) in the DL cells in vivo. (a) represents level of H2O2 in the DL cells. Values are mean ± SD and n ¼ 4.(b) represents level of total glutathione in the DL cells in vivo. Values are mean ± SD, where n ¼ 3. *p < 0.05; ***p < 0.001 (untreated vs. treated DL groups).

R.K. Koiri et al. / Biochimie 110 (2015) 52e61 59

H2O2 concentration was found to be remarkably increased (~6�) inthe DL cells from Ru(II)-CNEB treated DL mice (Fig. 5a). Thus, it isargued that Ru(II)-CNEB is able to modulate all the three enzymesof antioxidant pathway; SOD, catalase and GPx, to maintain higherlevel of H2O2 in the DL cells. Such a condition might drive the DLcells to undergo apoptosis, as increase in intracellular H2O2 isknown to cause a significant drop in cytosolic pH [51] which isconsidered accountable for translocation of Bax, a pro-apoptoticfactor, to mitochondria [49].

Fig. 6. Treatment with Ru(II)-CNEB (Rc) caused a significant decline in the expression of Bcland PARP1 cleavage (d) in the DL cells in vivo. a shows representative RT-PCR photograprepresent mean ± SD from 3 RT-PCR repeats. b shows representative western blot photograpmean ± SD from three western blot repeats. ***p < 0.001 (untreated vs. treated DL groups)

Though the mechanism by which Ru(II)-CNEB regulatesreciprocal changes in SODs vs catalase and GPx could be a matterof further investigation, in the present context, however,enhanced level of p53 protein (Fig. 2) could be considered as oneof the integrators of such enzymatic changes. This is because,not only a close association between p53 dependent modulationof ROS metabolism and cell apoptosis is on record [26,27] butalso, higher level of p53 protein has been found to enhance theexpression of pro-oxidant and proapoptotic factors [28,30].

2 with concomitantly increased expression of Bax (a and b), activation of caspase 9 (c)hs with the b-actin normalized densitometric ratio of Bcl2/Bax mRNA where valueshs with the b-actin normalized densitometric ratio of Bcl2/Bax where values represent.

R.K. Koiri et al. / Biochimie 110 (2015) 52e6160

Glutathione content is known to represent actual redox status ofthe cells and its enhanced level in the cancer cells is directlycorrelated with the tumor progression. Higher glutathione level, inaddition to providing multidrug and radiation resistance [52], hasbeen found to prevent apoptosis in the tumor cells [53]. Similarly,depletion of glutathione level has been reported to sensitize tumorcells to undergo apoptosis via release of cytochrome c [54]. Theanti-apoptotic role of Bcl-2 has also been linked with the gluta-thione content in the tumor cells [55]. We have shown previouslythat Ru(II)-CNEB causes release of mitochondrial cytochrome c [11]and according to Fig. 6a and b, it is evident to decline Bcl2/Bax ratioalso. Importantly these cellular alterations are consistent with asignificant decline in total glutathione (GSHþ GSSG) level in the DLcells (Fig. 5b) and thereby providing another biochemical mecha-nism by which Ru(II)-CNEB could be able to induce apoptosis in theDL cells.

One of the important aspects of p53 biochemistry is that it isconsidered to be involved in regulating apoptosis in the tumor cells[19,22,27,28]. Normally, p53 remains sequestered with Mdm2 inthe cytosol and thereby prevents cells to undergo apoptosis [56].This implies that release of p53 from Mdm2 could be one of themechanisms to induce apoptosis in the cells. This may happen dueto many cellular changes including increased DNA damage causedby the exogenous agents [57]. However, we observed that theincreasing concentration of Ru(II)-CNEB, when incubated in vitro at37 �C for 24 h with pBR322 plasmid DNA, it did not convertsupercoiled plasmid into the nicked circular DNA (Supplementarydata; Fig. S1) and thereby excluding direct nuclease activity ofthis compound. Therefore, in the present context, as reportedearlier [28,56], abrrent ellular signaling could be argued account-able for a significantly enhanced level of p53 in the DL cells fromRu(II)-CNEB treated DL mice (Fig. 2). Moreover, keeping aside themechanism by which p53 level gets enhanced in the DL cells, it isconsidered to act as a strong apoptotic inducer in multimodal ways.The enhanced cytoplasmic p53 level has been reported to activateBax and its oligomerization [58] which in turn, induces cytochromec release from the mitochondria [59]. Bcl2 is an anti-apoptoticfactor and therefore, declined Bcl2/Bax ratio is consideredaccountable for initiating intrinsic pathway of apoptosis [28]. In thepresent context, Ru(II)-CNEB significantly declined the level of Bcl2protein with a concomitant increase in Bax level (Fig. 6b). Thissuggested that Ru(II)-CNEB is able to reciprocally modulate level ofboth these factors resulting into a significant decline in Bcl2/Baxratio in the DL cells in vivo.

Caspase 9 activation is a hallmark of mitochondrial pathway ofapoptosis [60] and that of PARP-1 cleavage is associated with DNAfragmentation in the cells undergoing apoptosis [61]. We observedsignificant increase in the levels of cleaved caspase 9 and PARP-1 inthe DL cells from Ru(II)-CNEB treated DL mice (Fig. 6c and d).Consistent with these observations, Ru(II)-CNEBmediated release ofmitochondrial cytochrome c, the main initiator of the intrinsicpathway of apoptosis, in those DL cells, is already on record [11].Thus, taking together, these findings strongly advocate for p53mediated activation of Bcl2/Bax-cytochrome c release-caspase 9 ledintrinsic pathway of apoptosis in the DL cells due to treatment withRu(II)-CNEB in vivo. Though not much information is available oninduction of intrinsic pathway of apoptosis by metal complexes, aRu(II)-arene compound [Ru(g6-p-cymene)Cl2(pta)] has beendescribed to implicate this pathway for inducing apoptosis in theEhrlich ascite carcinoma [62]. Recently, another Ru(II)-complex(Ru(II) b-carboline complex) has also been demonstrated to induceapoptosis in the cancer cells by involving p53 [63]. In this context,the findings of this paper are of special merit with regard to eluci-dation of the biochemicalmechanismbywhich a Ru(II)-complex caninduce p53 dependent apoptosis in the tumor cells in vivo.

In conclusion, there is an evolving concept of restricting tumorgrowth by depriving tumor cells from adequate energy productionand by rendering them susceptible to oxidative stress. The presentarticle demonstrates that a Ru(II)-CNEB, characterized previously asan anti tumor compound, is evident to activate p53 mediatedglycolytic inhibition-oxidative stress-apoptosis pathway in the DLcells when administered in vivo. As such these findings provide abiochemical mechanism which can be utilized for defining phar-macological targets for the novel anticancer agents suitable forin vivo applications.

Conflict of interest

The authors declare no conflict of interest with respect to thisarticle.

Acknowledgements

This work was financially supported by a project from Depart-ment of Biotechnology (DBT), Govt. of India, (BT/PR5910/BRB/10/406/2005) sanctioned jointly to LM and SKT at BHU. The contri-bution of Dr. SK Dubey in synthesizing Ru(II)-CNEB in the lab of LMis also acknowledged. RKK thanks CSIR, Govt. of India for awardingSenior Research Fellowship during the tenure of this work at BHU.The authors are thankful to UGC Centre of Advanced Study pro-gramme to Department of Zoology and DBT-ISLS, BHU, forproviding facilities and assistance.

Appendix A. Supplementary data

Supplementary data related to this article can be found at http://dx.doi.org/10.1016/j.biochi.2014.12.021.

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